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Bedernjak, Alexandre (2010) Synthesis and Biological Evaluation of Novel

Chromogenic Substrates for the Enhanced Detection of Pathogenic Bacteria.

Doctoral thesis, University of Sunderland.

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SYNTHESIS AND BIOLOGICAL EVALUATION OF NOVEL

CHROMOGENIC SUBSTRATES FOR THE ENHANCED DETECTION

OF PATHOGENIC BACTERIA

Alexandre Bedernjak

A thesis submitted in partial fulfillment of the requirements of the University of

Sunderland for the degree of Doctor of Philosophy

This research programme was carried out in collaboration with the Freeman Hospital (Newcastle-upon-Tyne) and bioMérieux (La Balme-les-Grottes, France)

June 2010

L’important c’est pas la chute…

�

ACKNOWLEDGEMENTS

I would like to express the deepest and most sincere thanks to my supervisors

Prof. Roz Anderson and Prof. Paul Groundwater for their constant help, support,

encouragement, sharing of knowledge and infinite patience.

I also would like to thank my external supervisor Prof. John Perry for letting me

use his facilities at the Freeman Hospital and for his precious help with

microbiological testing.

Thanks to bioMérieux for financing this project, and thanks to the bioMérieux

team, Dr Sylvain Orenga, Céline Roger-Dalbert, and Marie Cellier for carrying the

microbiological testing.

Thanks also go to Prof. Arthur James for his advice and to his wife Gill for

welcoming us into their home in the Lake District for the most enjoyable team

meetings.

Special thanks go to Dr Andrey Zaytsev for his guidance, precious help and

advice during my first year.

Thanks to the chemiSpec team, Nicolas and Andrea, for their most precious

help with NMR spectroscopy, mass spectrometry and elemental analysis. Thanks to

the University of Sunderland technical staff, Arun, Jeff, Joan, Mick and Barry for their

help and letting me borrow equipment, glassware and chemicals. Thanks also to

Norman Turner for his most precious help in ordering chemicals.

Special thanks go to Rebecca for her precious help and support.

Thanks also to all my colleagues of lab 2.06 and lab 2.10, Adam, Alice, Gabi,

Linda, Lisa, Liz, Nagendra, Neil, Ning, Sam, Satya, Surresh, Pratap, Yu and Yang for

making these four years very enjoyable.

Merci aussi à Benjam et à Jérôme de s’être assuré que le retour au pays se

fasse toujours bien dans les règles de l’art.

Et finalement, un grand merci à mon père et à ma mère pour l’éducation qu’ils

m’ont donné, je me doute que ça ne devait pas être facile tous les jours, mais bon les

fruits d’un dur labeur sont là (ou presque).

ABSTRACT

The present work investigated the preparation of phenoxazinone derivatives

and evaluated their performances for the detection of pathogenic bacteria.

The first method investigated the condensation of nitroaminophenol with

tetrahalogenobenzoquinones; the corresponding nitrohalogenophenoxazinones were

all characterised and evaluated for the detection of nitroreductase activity in a range

of clinically important microorganisms. The detection of nitroreductase activity has

been previously suggested for the monitoring of bacterial growth; however,

nitrohalogenophenoxazinones were proven to be less suitable for this purpose than

other, previously reported, nitroreductase substrates.

The second route investigated the synthesis of phenoxazinone derivatives via

the oxidative cyclisation of diamino-dihydroxydiphenylethers and of

diaminobenzoquinones. The reactive intermediates were trapped and characterised

in order to rationalize the mechanism of formation of aminophenoxazinones via this

route. 7- and 8-Aminophenoxazinones derivatives were prepared and further

derivatised with β-alanine. Similarly, some nitrohalogenophenoxazinones were

reduced to their corresponding aminophenoxazinones and derivatised with β-alanine.

Thirteen new chromogenic substrates were prepared, characterised and

evaluated for their sensitivity to detect β-alanine aminopeptidase on agar medium;

this enzyme is expressed by only three types of bacteria, the most important being

Pseudomonas aeruginosa, a pathogen commonly known to affect cystic fibrosis

sufferers. Their performance for the detection of Pseudomonas aeruginosa were

compared to the lead compound (7-N-(β-alanyl)amino-1-pentylphenoxazin-3-one),

the substrate contained in a commercially available medium, chromIDTM ID Ps.

aeruginosa. The substrates, if hydrolysed, resulted in a low colouration of the

colonies when compared to the lead compound; however, 2-pentyl substituted

aminophenoxazinones were found to be less toxic and had an excellent affinity for

the bacterial colonies.

.

SYMBOLS AND ABBREVIATIONS

calc. calculated

cat. catalytic

conc. concentrated

DBU 1,8-diazabicyclo[5-4-0]undec-7-ene

d doublet

dd doublet of doublets

ddd doublet of doublet of doublets

dt doublet of triplets

DCM dichloromethane

DEPT Distortionless Enhancement by Polarization Transfer

DNA deoxyribonucleic acid

DME dimethoxyethane

DMF dimethylformamide

DMS dimethylsulfate

DMSO dimethylsulfoxide

Et2O diethyl ether

EtOAc ethyl acetate

EtOH ethanol

Et3N triethylamine

h hour

[H] reduction

HMBC Heteronuclear Multiple Bond Correlation

HMQC Heteronuclear Multiple Quantum Coherence

Hz hertz

IBCF iso-butylchloroformate

J coupling constant

M molarity

M + positive molecular ion

M - negative molecular ion

MeOH methanol

min minute

mL millilitre

mol mole

mmol mmole

mg milligram

m.p. melting point

MMPP magnesium monoperoxyphthalate

MRSA meticillin-resistant Staphylococcus aureus

MW molecular weight

m/Z mass over charge of the ion

n-BuLi n-butyl lithium

NMM N-methylmorpholine

NMR nuclear magnetic resonance

NMP N-methyl-2-pyrrolidone

[O] oxidation

PCR polymerase chain reaction

ppm parts per million

q quartet

quat. quaternary

RNA ribonucleic acid

s singlet

sex. sextuplet

T temperature

t triplet

td triplet of doublets tBoc tert-butoxycarbonyl

TFA trifluoroacetic acid

THF tetrahydrofuran

TLC thin layer chromatography

U.V. ultra-violet

v/v volumic percentage

w/w weight percentage

δ NMR chemical shift

∆ heat

ε molar extinction coefficient

TABLE OF CONTENTS

CHAPTER ONE: INTRODUCTION

1.1 Bacterial detection 1

1.1.1 Classical detection method 2

1.1.1.1 The Gram stain 2

1.1.2 New methods of identification 5

1.1.2.1 Polymerase chain reaction (PCR) 5

1.1.2.2 Bacteriophages 6

1.1.2.3 Enzyme-linked immunosorbent assay (ELISA) 8

1.1.2.4 Chromogenic media 9

1.1.2.4.1 Agar-based media 9

1.1.2.4.2 Micro gallery and automated systems 10

1.1.3 Chromogenic and fluorogenic dyes 11

1.1.3.1 DNA fluorescent dyes 11

1.1.3.2 pH indicators 12

1.1.3.3 Redox indicators 13

1.1.3.4 Enzyme substrates 14

1.1.3.4.1 Simultaneous capture chromogenic substrates 15

1.1.3.4.2 Post-incubation coupling chromogenic substrates 19

1.1.3.4.3 Intramolecular rearrangement/electron conjugation

chromogenic substrates 21

1.1.3.4.4 Self coloured chromogenic/fluorogenic substrates 23

1.1.3.4.4.1 Fluorogen 24

1.1.3.4.4.2 Chromogens 25

1.1.4 Enzyme targets 27

1.1.4.1 Esterases and lipases 27

1.1.4.2 Glycosidases 28

1.1.4.2.1 β-D-Glucuronidase 28

1.1.4.2.2 β-D-Galactosidase 29

1.1.4.2.3 α-D-Galactosidase 30

1.1.4.2.4 β-D-Glucosidase 30

1.1.4.2.5 α-D-Glucosidase 31

1.1.4.2.6 β-Hexoaminidase 31

1.1.4.3 Phosphatase 32

1.1.4.3.1 Phosphatidylinositol phospholipase C (PI-PLC) 33

1.1.4.4 Peptidase 33

1.1.4.4.1 Pyroglutamyl aminopeptidase 35

1.1.4.4.2 L-Alanine aminopeptidase 35

1.1.4.4.3 β-Alanine aminopeptidase 36

1.1.4.5 Nitroreductase enzymes 37

1.2 Occurrence of the phenoxazinone core 38

1.2.1 Natural products 38

1.2.1.1 Ommochromes 38

1.2.1.2 Mould metabolites 40

1.2.1.3 Actinomycins 41

1.2.1.4 Biosynthesis of the phenoxazinone core 41

1.2.2 Phenoxazinone dyes 44

1.2.2.1 Litmus and orceins 44

1.3 Chemical synthesis of the phenoxazinone core 46

1.3.1 Oxidative condensation of o-aminophenol 46

1.3.2 Oxidative mixed condensation of o-aminophenol 48

1.3.3 Condensation of o-aminophenols with hydroxyquinones 49

1.3.4 Condensation of o-aminophenol with quinolin-5,8-dione 50

1.3.5 Condensation of o-aminophenols with halogenated quinones 51

1.3.6 Condensation of p-nitrosoaniline and o-nitrosophenols

with phenols 52

1.3.7 Condensation of benzoquinonechlorodiimide with resorcinol 54

1.3.8 Reductive cyclisation of 5-hydroxy-2'-nitrodiphenyl ethers 55

1.3.9 Oxidative cyclisation of 2,5-dinitro-2',5'-dihydroxy-

diphenylether 56

1.3.10 Oxidation of phenoxazines 58

1.4 Aims 59

CHAPTER TWO: SYNTHESIS OF PHENOXAZINONE SUBSTRATES

2.1 Synthesis of halogenophenoxazin-3-one substrates 60

2.1.1 Preparation of nitro-1,2,4-trihalogenophenoxazin-3-ones

130αααα-γγγγa-c 60

2.1.1.1 Nitro-1,2,4-trifluorophenoxazin-3-ones 130αααα-γγγγa 63

2.1.2 Synthesis of 8-amino-1,2,4-trihalogenophenoxazin-3-ones

165a-c 67

2.2 Synthesis of phenoxazinones via oxidative cyclisation 68

2.2.1 Previous work 68

2.2.2 Substitution strategy 69

2.2.3 Retrosynthetic analysis 70

2.2.4 Synthesis of 2,5-dimethoxybenzaldehyde 74

2.2.4.1 From hydroquinones 173a-c 74

2.2.4.2 From 2-hydroxy-5-methoxybenzaldehyde 182 74

2.2.4.2.1 Sonogashira cross-coupling 76

2.2.4.2.2 Suzuki cross-coupling 79

2.2.4.3 From 1,4-dimethoxy-2-bromobenzene 174 81

2.2.5 Synthesis of 2,5-dimethoxyphenols 150a-f 83

2.2.6 Bromination of 3,5-dimethoxyphenol 177 85

2.2.7 Oxidation of olivetol 181 85

2.2.8 Synthesis of diphenylethers 169a-g, 175b-c and 152a-b 87

2.2.8.1 Reaction with 2,4-dinitro-1-fluorobenzene 168 87

2.2.8.2 Reaction with 2,5-dinitro-1-fluorobenzene 151 90

2.2.9 Attempted substitution of diphenylether 92

2.2.9.1 From 2,4-dinitro-3'-bromo-2',5'-dimethoxy-diphenylether 92

2.2.9.2 From 2,4-dinitro-2',5'-dimethoxy-diphenylether 169g 94

2.2.10 Deprotection of diphenylethers 169b,g and 227b with BBr3 95

2.2.11 Oxidative demethylation of diphenylethers 169a,c,d,f

and 152c-d using of cerium (IV) ammonium nitrate 97

2.2.12 Synthesis of 7- and 8-aminophenoxazin-3-ones 101

2.2.12.1 From 2,4-dinitro-2',5'-dihydroxydiphenylethers 170a-b 101

2.2.12.2 From 2,4 and 2,5-dinitrophenoxy-2',5'-benzoquinones 103

2.2.14 Trapping of reactive species 105

2.2.14.1 2,4-Diamino-2',5'-dihydroxydiphenylether 220a 105

2.2.14.2 2,4-Diaminophenoxybenzoquinones 171b,c,e 106

2.2.14.3 2,4-Diamino-2',5'-dihydroxydiphenylether 154a 109

2.2.15 Mechanism of cyclisation 110

2.3 Synthesis of β-alanine substrates 235a-b and 237a-i 112

2.3.1 From 7-aminophenoxazin-3-ones 59d-e 112

2.3.2 From 8-aminophenoxazin-3-ones 165a-c and 167a-f 115

2.3.2.1 N-β-alanine derivatives 115

2.3.2.2 N- and O- di-β-alanine derivatives 117

2.4 U.V.-Visible absorption 120

2.4.1 Comparison of 7- and 8-aminophenoxazin-3-ones 59d-e

and 167d-e 120

2.4.2 Quenching effect of ββββ-alanine for substrates 235a-b and 237a-c 123

CHAPTER THREE: MICROBIOLOGICAL RESULTS

3.1 Nitroreductase activity 126

3.1.1 Results and discussion 127

3.2 Testing of β-alanyl halogenated phenoxazinone derivatives

237g-i and 240a-b 135

3.2.1 Toxicity screening 136


3.3 Testing of β-alanylalkylphenoxazin-3-one derivatives

237a-g and 235a-b 141


3.3.2.1 Lipophilicity 148

3.4 Conclusion 149

CHAPTER FOUR: CONCLUSION

4.1 Conclusion and future work 151

4.1.1 Halogenophenoxazinone substrates 151

4.1.1.1 Summary 151

4.1.1.2 Future work 153

4.1.2 Alkylphenoxazinone substrates 154

4.1.2.1 Summary 154

4.1.2.2 Future work 158

CHAPTER FIVE: EXPERIMENTAL

5.1 General experimental 161

5.2 Synthesis 161

5.2.1 General procedure for the preparation of

nitro-1,2,4-trihalegeno-3H-phenoxazin-3-one 130αααα-γγγγ a-c 161

5.2.1.1 7-Nitro-1,2,4-trifluoro-3H-phenoxazin-3-one 130ααααa 162

5.2.1.2 7-Nitro-1,2,4-trichloro-3H-phenoxazin-3-one 130ααααb 162

5.2.1.3 7-Nitro-1,2,4-tribromo-3H-phenoxazin-3-one 130ααααc 163

5.2.1.4 8-Nitro-1,2,4-trifluoro-3H-phenoxazin-3-one 130ββββa 163

5.2.1.5 8-Nitro-1,2,4-trichloro-3H-phenoxazin-3-one 130ββββb 165

5.2.1.6 8-Nitro-1,2,4-tribromo-3H-phenoxazin-3-one 130ββββc 165

5.2.1.7 9-Nitro-1,2,4-trifluoro-3H-phenoxazin-3-one 130γγγγa 166

5.2.1.8 9-Nitro-1,2,4-trichloro-3H-phenoxazin-3-one 130γγγγb 166


8-amino-1,2,4-trihalegeno-3H-phenoxazin-3-one 165a-c 167

5.2.2.1 8-Amino-1,2,4-trifluoro-3H-phenoxazin-3-one 165a 167


8-N-(N'-tbutoxycarbonyl-β-alanyl)amino-1,2,4-trihalogeno-

3H-phenoxazin-3-ones 236g-i 168

5.2.3.1 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2,4-trifluoro-3H-

phenoxazin-3-one 236g 169

5.2.3.2 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2,4-trichloro-3H-

phenoxazin-3-one 236h 170

5.2.3.3 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2,4-tribromo-3H-

phenoxazin-3-one 236i 171

5.2.4 General procedure for the deprotection of 8-N-(N'-tbutoxycarbonyl-

β-alanyl)amino-1,2,4-halogeno-3H-phenoxazin-3-one 237g-i 172

5.2.4.1 8-N-(β-Alanyl)amino-1,2,4-trifluoro-3H-phenoxazin-3-one

TFA salt 237g 172

5.2.4.2 8-N-(β-Alanyl)amino-1,2,4-trichloro-3H-phenoxazin-3-one

TFA salt 237h 173

5.2.4.3 8-N-(β-Alanyl)amino-1,2,4-tribromo-3H-phenoxazin-3-one

TFA salt 237i 173


8-N-(N'-tbutoxycarbonyl-β-alanyl)amino-1,2,4-trihalogeno-10H-

phenoxazin-3-yl 3''-(N-tbutoxycarbonyl-amino)propanoate

239a-b 174

5.2.5.1 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2,4-trifluoro-10H-

phenoxazin-3-yl 3''-(N''-tbutoxycarbonylamino)propanoate 239a 175

5.2.5.2 8-N-(N-tButoxycarbonyl-β-alanyl)amino-1,2,4-trichloro-10H-

phenoxazin-3-yl-3''-(N-tbutoxycarbonylamino)propanoate 239b 175

5.2.6 General procedure for the deprotection of 8-N-(N-tbutoxycarbonyl-

β-alanyl)amino-1,2,4-trihalogeno-10H-phenoxazin-3-yl-

3''-(N-tbutoxycarbonyl-amino)propanoate 240a-b 176

5.2.6.1 3-O-(8-N-(β-Alanyl)amino-1,2,4-trifluoro-10H-phenoxazin-3-yloxy)-

1''-oxopropane-3''-aminium ditrifluoroacetate salt 240a 177

5.2.6.2 3-O-(8-N-(β-Alanyl)amino-1,2,4-trichloro-10H-phenoxazin-3-yloxy)-

1''-oxopropane-3''-aminium ditrifluoroacetate salt 240b 177


dimethoxybenzenes 172a-c 178

5.2.7.1 1,4-Dimethoxy-2,3-dimethylbenzene 172a 179

5.2.7.2 1,4-Dimethoxy-2,3,5-trimethylbenzene 172b 179

5.2.7.3 1,4-Dimethoxy-2-tbutylbenzene 172c 180

5.2.8 Procedure for the lithiation of

2-bromo-1,4-dimethoxybenzene 178 180

5.2.8.1 1,4-Dimethoxy-2-pentylbenzene 196 180

5.2.9 General procedure for the formylation of substituted

dimethoxybenzene 172a-c, 196 and 189 via the Duff reaction 181

5.2.9.1 2,5-Dimethoxy-3,4-dimethylbenzaldehyde 149a 182

5.2.9.2 2,5-Dimethoxy-3,4,6-trimethylbenzaldehyde 149b 182

5.2.9.3 2,5-Dimethoxy-4-tbutylbenzaldehyde 149c 183

5.2.9.4 2,5-Dimethoxy-4-pentylbenzaldehyde 197 183

5.2.9.5 2,4-Dinitro-2',5'-dimethoxy-4'-carbaldehydediphenylether 211 184

5.2.10 Halogenation of 2-hydroxy-5-methoxybenzaldehyde 182 185

5.2.10.1 3-Bromo-2-hydroxy-5-methoxybenzaldehyde 183 185

5.2.10.2 3-Iodo-2-hydroxy-5-methoxybenzaldehyde 186a and

2,2'-dihydroxy-5,5'-dimethoxybiphenyl-3,3'-dicarboxaldehyde

186b 186

5.2.11 Protection of 3-halogeno-2-hydroxy-5-methoxybenzaldehyde 187

5.2.11.1 3-Bromo-2,5-dimethoxybenzaldehyde 178 and

2,5-dimethoxybenzaldehyde 149 187

5.2.11.2 3-Iodo-2,5-dimethoxybenzaldehyde 185 188

5.2.12 Sonogoshira cross-couping 189

5.2.12.1 Preparation of 2,5-dimethoxy-1-pentyn-1'-ylbenzaldehyde 184 189

5.2.13 Suzuki cross-coupling 191

5.2.13.1 Preparation of 2,5-dimethoxybiphenyl-3-carbaldehyde 194 191

5.2.13.2 Attempted preparation of 3-(2',4'-dinitrophenoxy)-

2,5-dimethoxybiphenyl 169f 191

5.2.14 General procedure for the Baeyer-Villiger oxidation

of benzaldehydes 149a-d, 178, 194 and 197 192

5.2.14.1 2,5-Dimethoxyphenol 150g 193

5.2.14.2 2,5-Dimethoxy-3,4-dimethylphenol 150a 194

5.2.14.3 2,5-Dimethoxy-3,4,6-trimethylphenol 150b 194

5.2.14.4 2,5-dimethoxy-4-tbutylphenol 150c 195

5.2.14.5 2,5-Dimethoxy-4-pentylphenol 150d 197

5.2.14.6 3-Bromo-2,5-dimethoxyphenol 150e 197

5.2.14.7 2,5-Dimethoxy-biphenyl-3-ol 150f 198

5.2.15 Bromination 198

5.2.15.1 2,6-Dibromo-3,5-dimethoxyphenol 176 198

5.2.15.1 2,4,6-Tribromoolivetol 205 199

5.2.16 Oxidation of tribromoolivetol 205: 2,6-dibromo-3-hydroxy-

5-pentyl-1,4-benzoquinone 206 200

5.2.17 4-Nitro-2-fluoroaniline 210a and 6-nitro-2-fluoroaniline 210b 201

5.2.18 General procedure for the oxidation of fluoroanilines 202

5.2.18.1 2,5-Dinitro-1-fluorobenzene 151 203

5.2.19 General procedure for the preparation of biarylethers

169a-g and 152c-d 203

5.2.19.1 2,4-Dinitro-2',5'-dimethoxydiphenylether 169g 204

5.2.19.2 2,4-Dinitro-2',5'-dimethoxy-3',4'-dimethyldiphenylether 169a 205

5.2.19.3 2,4-Dinitro-2',5'-dimethoxy-3',4',6'-trimethyldiphenylether 169b 205

5.2.19.4 2,4-Dinitro-2',5'-dimethoxy-4'-tbutyldiphenylether 169c 206

5.2.19.5 2,4-Dinitro-2',5'-dimethoxy-3'-pentyldiphenylether 169d 207

5.2.19.6 2,4-Dinitro-6'-bromo-2',5'-dimethoxydiphenylether 169e 207

5.2.19.7 3-(2',4'-Dinitrophenoxy)-2,5-dimethoxybiphenyl 169f 208

5.2.19.8 2,4-Dinitro-3',5'-dimethoxydiphenylether 175c 209

5.2.19.9 2,4-Dinitro-2',6'-dibromo-3',5'-dimethoxydiphenylether 175b 209

5.2.19.10 2,5-Dinitro-4'-tbutyl-2',5'-dimethoxydiphenylether 152c 210

5.2.19.11 2,5-Dinitro-2',5'-dimethoxy-3'-pentyldiphenylether 152d 210

5.2.19.12 Attempted preparation of 2,4-dinitro-3'-pentyl-

2',5'-dimethoxydiphenylether 169p 211

5.2.20 General procedure for the deprotection of substituted

2,4-dinitro-2',5'-dimethoxydiphenyl ethers 169b and 169g 212

5.2.20.1 2,4-Dinitro-2',5'-dihydroxydiphenylether 170a 213

5.2.20.2 2,4-Dinitro-2',5'-dihydroxy-3',4',6'-trimethyldiphenylether 170b

and 2,4-dinitrophenoxy-3',4',6'-trimethyl-1',4'-benzoquinone 179b 213

5.2.20.3 2,4-dinitro-2',6'-dibromo-3'-hydroxy-5'-methoxydiphenylether

212 and 2,4-dinitro-2',6'-dibromo-3',5'-dihydroxydiphenylether

214 214

5.2.21 General procedure for the oxidation of the substituted

2,4-dinitro-2',5'-dimethoxydiphenylether 169a,c,d,f and

2,4-dinitro-2',5'-dimethoxydiphenylether 219a-b 215

5.2.21.2 2,4-Dinitrophenoxy-3',4'-dimethyl-2',5'-benzoquinone 179a 216

5.2.21.3 2,4-Dinitrophenoxy-4'-tbutyl-2',5'-benzoquinone 179c 216

5.2.21.4 2,4-Dinitrophenoxy-3'-phenyl-2',5'-benzoquinone 179d 217

5.2.21.5 2,4-Dinitrophenoxy-4'-pentyl-2',5'-benzoquinone 179e 218

5.2.21.6 2,5-Dinitrophenoxy-4'-t butyl-2',5'-benzoquinone 219a 218

5.2.21.7 2,5-Dinitrophenoxy-4'-pentyl-2',5'-benzoquinone 219b 219

5.2.22 General procedure for the trapping of 2,4-diamino-

2',5'-dihydroxydiphenylether 220a and diaminophenoxy-

2',5'-benzoquinone 171a,c,d and 154a 220

5.2.22.1 2,4-Diacetamido-2',5'-dihydroxydiphenylether 222 220

5.2.22.2 2,4-Diacetamido-3',4'-dimethyl-2',5'-benzoquinone 223a 221

5.2.22.3 2,4-Diacetamido-4'-tbutyl-2',5'-benzoquinone 223b 222

5.2.22.4 2,4-Diacetamido-4'-pentyl-2',5'-benzoquinone 223c 222

5.2.22.5 2,5-Diacetamido-2',5'-dihydroxy-4'-tbutyldiphenylether 258 223

5.2.22.6 2-Acetoxy-5-acetamido-1-phenylamino-3',4'-dimethyl-2',5'-

benzoquinone 224 224

5.2.23 General procedure for the preparation of 8-amino-alkyl-3H-

phenoxazin-3-ones 167a-e and 7-amino-2-alkyl-3H-

phenoxazin-3-ones 59d-e 225

5.2.23.1 8-Amino-3H-phenoxazin-3-one 167a 226

5.2.23.2 8-Amino-1,2-dimethyl-3H-phenoxazin-3-one 167b 227

5.2.23.3 8-Amino-1,2,4-trimethyl-3H-phenoxazin-3-one 167c 227

5.2.23.4 8-Amino-2-tbutyl-3H-phenoxazin-3-one 167d 228

5.2.23.5 8-Amino-2-pentyl-3H-phenoxazin-3-one 167e 228

5.2.23.6 8-Amino-1-phenyl-3H-phenoxazin-3-one 167f 229

5.2.23.7 7-Amino-2-tbutyl-3H-phenoxazin-3-one 59d 230

5.2.23.8 7-Amino-2-pentyl-3H-phenoxazin-3-one 59e 230

5.2.24 General procedure for the preparation of 7-N and

8-N-(N'-tbutoxycarbonyl-β-alanyl)amino-3H-phenoxazin-3-ones

234a-b and 236a-f 231

5.2.24.1 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-3H-phenoxazin-3-one

236a 232

5.2.24.2 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2-dimethyl-3H-

phenoxazin-3-one 236b 232

5.2.24.3 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1,2,4-trimethyl-3H-

phenoxazin-3-one 236c 233

5.2.24.4 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-2-tbutyl-3H-

phenoxazin-3-one 236d 234

5.2.24.5 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-2-pentyl-3H-

phenoxazin-3-one 236e 234

5.2.24.6 8-N-(N'-tButoxycarbonyl-β-alanyl)amino-1-phenyl-3H-

phenoxazin-3-one 236f 235

5.2.24.7 7-N-(N'-tButoxycarbonyl-β-alanyl)amino-2-tbutyl-3H-

phenoxazin-3-one 234a 236

5.2.24.8 7-N-(N'-tButoxycarbonyl-β-alanyl)amino-2-pentyl-3H-

phenoxazin-3-one 234b 236

5.2.25 General procedure for the deprotection of 8-N-(N'-tbutoxycarbonyl-

β-alanyl)amino-alkyl-3H-phenoxazin-3-one 236a-f and

7-N-(N'-tbutoxycarbonyl-β-alanyl)amino-alkyl-3H-phenoxazin-3-one

268a-b 237

5.2.25.1 8-N-(β-Alanyl)amino-3H-phenoxazin-3-one TFA salt 237a 238

5.2.25.2 8-N-(β-Alanyl)amino-1,2-dimethyl-3H-phenoxazin-3-one

TFA salt 237b 238

5.2.25.3 8-N-(β-Alanyl)amino-1,2,4-trimethyl-3H-phenoxazin-3-one

TFA salt 237c 239

5.2.25.4 8-N-(β-Alanyl)amino-2-tbutyl-3H-phenoxazin-3-one

TFA salt 237d 239

5.2.25.5 8-N-(β-alanyl)amino-2-pentyl-3H-phenoxazin-3-one

TFA salt 237e 240

5.2.25.6 8-N-(β-Alanyl)amino-1-phenyl-3H-phenoxazin-3-one

TFA salt 237f 241

5.2.25.7 7-N-(β-Alanyl)amino-2-tbutyl-3H-phenoxazin-3-one TFA 235a 241

5.2.25.8 7-N-(β-Alanyl)amino-2-pentyl-3H-phenoxazin-3-one

TFA salt 235b 242

5.3 Microbiological testing 243

5.3.1 Preparation of the medium 243

5.3.1.1 Nitrophenoxazinones substrates 243

5.3.1.2 Trihalogenophenoxazin-3-one substrates 237g-i and 240a-b 244

5.3.1.3 Alkylphenoxazin-3-one substrates 237a-f and 235a-b 245

5.3.2 Inoculation 245

5.3.2.1 Individual inoculation 245

5.3.2.2 Multipoint inoculation 246

CHAPTER SIX: REFERENCES

6.1 References 248

APPENDIX

CHAPTER ONE:

INTRODUCTION

CHAPTER ONE INTRODUCTION

1

Microorganisms are ubiquitous in our environment: they colonise and adapt to the

most extreme environment (extremophiles) or live in symbiosis with other organisms,

such as the human commensal bacteria Staphylococci (skin, 2001JID170),

Streptococci (saliva and teeth, 1996AOB1133) or Bacteroid, Eubacterium and

Bifidobacterium (colon, 2005CPD1047) to name a few. If the presence of such

bacteria is well tolerated by, and even necessary to, humans (nutrients; immune

system stimulation), the proliferation of exogenous pathogenic bacteria can lead to

life-threatening conditions.

The presence of pathogenic bacteria can have disastrous consequences in hospitals,

food and water, to name three highly significant areas of our health and well-being.

Their identification in such domains is necessary and has become a corner stone of

such areas.

The present work focuses on the development of specific chromogenic enzyme

substrates allowing for the rapid detection of nosocomial bacteria. Identification using

chromogenic enzyme substrates has acquired a significant impact for the detection of

human pathogens. Their simplicity and reliability, shorter identification process and

fewer complementary tests required, respond to the need for quick results in a

hospital context. Indeed, the successful treatment and eradication of a pathogen rely

on its quick and correct identification.

Many methods have been developed, and are still under development, for the

accurate and rapid detection of microorganisms (1996MI455, 2000MI549). Full

details of each method are out of the scope of the present work and only a brief

overview of the existing and most relevant techniques will be given here. The

description of techniques using fluorogenic and, more particularly, chromogenic

enzyme substrates as a means of detection will be more carefully detailed with an

insight into their properties, structures and mode of action. The second part of this

introduction will be dedicated to the occurrence and preparation of the phenoxazin-3-

one core, which is the core of the new chromogenic substrates prepared herein.

1.1 Bacterial detection

Identification of a bacterium requires quantifying a relationship based on its

morphology, physiology and chemical structure (B-1993MI01). Bacteria are

characterized, defined and named after the appearance of their colonies, the


2

morphology of a single cell and respective arrangement of several cells, their growth

characteristics, their biochemistry and their reaction to inhibitory tests, the need for

specific nutrients or conditions for growth, their reaction to antiserum and their

genetics. The detection of a bacterium involves a series of tests allowing the

observation of a determinant character allowing its relation to a previously described

genus and species. It is generally admitted that no single test provides a definitive

identification of an unknown microorganism. The traditional identification methods

involve culture of the bacteria for enrichment and isolation of the subsequent

bacterial colonies, followed by screening using a series of biochemical tests and

serological confirmation (1990MI497).

1.1.1 Classical detection methods

1.1.1.1 The Gram stain

The Gram stain, developed in 1884 by Christian Gram, was the first taxonomic test of

its kind allowing such an important differentiation within the bacterial kingdom. The

Gram stain procedure (1884MI185) involves the use of an aqueous solution of

gentian violet (crystal violet, 1), along with a saturated

solution of iodine and potassium iodide as mordant.

Decolorisation of the bacterial cell is then attempted by

washing with ethanol or acetone, leaving Gram-positive

bacteria stained in violet. A counter stain of carbol fuschin

2 is finally applied, dying any Gram-negative cells in red.

Several factors can influence the outcome of the Gram

stain, such as the age of the culture and the composition

of the cultivation medium (1990JAB822). Errors in

determining the Gram reaction have been observed, with bacteria, known as Gram

variable, showing a variable behaviour toward the Gram stain.

The ability of Gram-positive organisms to retain the dye more effectively than Gram-

negative organisms when exposed to a solvent is related to major differences within

the structure of the bacterial cell wall. Gram-positive bacterial cell walls (Figure 1.1)

consist of a thick homogeneous layer of peptidoglycan, covalently bound to linear

anionic polymeric teichoic acids (ribitol and glycerol units linked by phosphodiesters,

2002MI46S), situated outside the plasma membrane.

1, R1 = CH3, R2 = H 2, R1 = H, R2 = CH3

N

NN

R1

R1

R1R1

R1

R1

Cl

R2


3

Figure 1.1: Schematic and simplified structure of Gram-positive (left) and Gram-negative (right) bacterial cell wall (adapted from B-2008MI02).

Peptidoglycan layer

Plasma membrane

Outer membrane

Periplasmic space

Periplasmic space

Protein

Porin

Braun’s lipoprotein

: N-Acetylglucosamine (NAG)

: N-Acetylmuramic acid (NAM)

: ribitol

: glycoside unit

: glycerol

: phosphate

Techoic acid

Polysaccharides O side chain


4

Peptidoglycan is a mesh-like polymer composed of covalently bound N-

acetylglucosamine (NAG) and N-acetyl muramic acid (NAM) subunits.

The peptidoglycan layer is highly crosslinked in Gram positive organisms;

crosslinking occurs via the carboxylic acid residues of NAM. With Gram-positive

bacteria, crystal violet is thought to be retained behind the peptidoglycan layer, which

pores are being shrunk upon the decolorisation process.

The Gram-negative bacterial cell wall is a relatively more complex structure (Figure

1.1). It consists of a thin, poorly crosslinked peptidoglycan layer recovered by an

outer membrane; the coherence of this structure is ensured by Braun’s lipoproteins

(Figure 1.1). The outer membrane, which is the main self-protective structure of

Gram-positive bacteria against toxic compounds, is composed of three main

elements: the phospholipid layer, the lipopolysaccharide layer and the

polysaccharide O-side chain. This highly lipophilic external structure exercises a

control over the permeation of hydrophilic compounds, whereas the porin channels it

contains (Figure 1.1) allow the permeation of small hydrophilic molecules such as

nutrients (2001MI215).

With Gram-negative bacteria, the thin peptidoglycan layer and the outer membrane,

which permeability is affected by ethanol, fail to retain crystal violet upon the

decolorisation process, leaving the cells colourless.

Other, yet less popular, dye-based Gram differentiation methods have since been

developed. The growth of bacteria on medium containing 8-

anilino-1-naphthalene sulphonic acid (ANS, 3), a hydrophobic

fluorescent dye, was found to differentiate Gram-negative

bacteria from Gram-positive by selective fluorescence. The

hydrophobic interactions occurring upon adsorption of the dye

onto the bacterial proteins, present in the lipopolysaccharide layer of the Gram-

negative outer membrane, result in a dramatic increase of ANS quantum yield

(1980AEM372). The lack of interaction with the outer peptidoglycan layer of Gram-

negative bacteria left the latter non-fluorescent, the intrinsic quantum yield of ANS

being very low in aqueous environment.

More recently, Mason et al. reported a live Gram stain

method using two fluorescent nucleic acid binding dyes,

hexidium iodide (HI, 4) and SYTO 13 (1998AEM2681).

NH SO3H

3

NH2N

NH2

I

4, λex/em = 518nm/600nm


5

While Gram-positive bacteria did permeate both HI and SYTO 13, Gram-negative

only permeated SYTO 13. This resulted in the observation of an orange fluorescence

for Gram-positive bacteria, as the simultaneous presence of both dyes resulted in a

quenching of SYTO 13 fluorescence by HI (1998AEM2681), and a green

fluorescence for Gram-negative bacteria. The outer lipopolysaccharide layer is

thought to be responsible for the exclusion of HI (1998AEM2681).

1.1.2 New methods of identification

1.1.2.1 Polymerase chain reaction (PCR)

The polymerase chain reaction (PCR) is defined as “a primer-mediated enzymatic

amplification of specifically cloned or genomic DNA sequence” (B-2002MI03). The

PCR process involves denaturation of a DNA sequence, hybridization of the primers

(short oligonucleotides sequences) and polymerization of the desired sequence by a

thermostable DNA polymerase, Taq (from the thermophile Thermus aquaticus).

These three successive steps, which constitute a cycle, are realized in a thermal

cycler, allowing each step to take place at a different, optimum temperature: 95°C

(denaturation), 60°C (primer hybridization) and 72°C (polymerization). This cycle is

typically repeated 20 to 40 times, at which point billions of copies of the gene

sequence are available (number of cloned sequence = 2n cycles). Interpretation of the

results is done by staining the PCR product with a phenantridium dye (analogous to

4), followed by separation using gel electrophoresis. The recent emergence of real-

time PCR can by-pass this interpretation step by

monitoring the increase of fluorescence during the

amplification process using double-stranded DNA

specific dyes such as SYBR Green I 5.

The ability to replicate a unique gene sequence has

proven to be highly useful in the identification of

pathogenic bacteria and PCR is becoming increasingly

important as a rapid means of detection (1998JCM2810).

Virtually any species-specific nucleotide sequence can be selected for amplification,

but replication of gene sequence determinants of the pathogenic character of a

bacterium is sometimes key to identification: the vtx1 and vtx2 genes encoding for

the production of verotoxin, responsible for the hemorrhagic colitis linked to E. coli

O157 : H7 infection (2010MI7), can easily discriminate E. coli O157 : H7 from non

N

N

S

N

N

5, λex/em = 494nm/521nm


6

virulent E. coli strains. The mecA gene, encoding for the production of penicillin-

binding protein 2a (1997CMR781), is another example of useful gene marker which

can discriminate meticillin-resistant Staphylococcus aureus (MRSA) from non-

resistant Staph. aureus species.

The concomitant use of several pairs of primers (multiplex PCR) has been

successfully applied to the simultaneous detection of some nosocomial pathogens,

such as Staphylococcus aureus, Staphylococcus epidermidis, Pseudomonas

aeruginosa, Acinetobacter baumanii, and Klebsiella pneumonia (2009JMM329), and

common food contaminants, Salmonella spp. (2009MI348, 2009MI43), Listeria

monocytogenes (2006MI763) and Campylobacter jejuni (2003AEM1383).

1.1.2.2 Bacteriophages

Bacteriophages or phages are viruses that infect bacteria cells only. They can

recognize and bind to specific receptors on the outer membrane of bacterial cells, i.e.

an amino acid sequence of a surface protein. The high specifity of bacteriophages for

their respective host has led to the development of an original approach to bacterial

detection, which was first demonstrated by Ulitzur and Kuhn (B-1987MI04).

For the purpose of pathogen detection, phages are genetically engineered to include

a reporter gene, the majority being bioluminescence genes (lux) from naturally

bioluminescent species, such as Vibrio fischeri. Upon infection of the bacterium by

the genetically modified phage, the lux reporter gene is introduced along with the

phage DNA into the bacterial host and then expressed (Scheme 1.1). As the lux

gene encodes for the production of luciferase (2002FRI863), the infected bacterium

becomes bioluminescent. The production of bacterial bioluminescence results from

the aerobic luciferase-catalysed oxidation of an aliphatic aldehyde (e.g. dodecanal 6)

in the presence of reduced flavin mononucleotide (FMNH2) (Scheme 1.1,

1991MR123). Monitoring of the resulting light emission is therefore indicative of the

presence of the bacterium.

Assays using luciferase reporter phages have been reported for the detection of the

following pathogens: E. Coli O157 : H7 (1996MI152), Listeria monocytogenes

(1997AEM2961), Mycobacterium tuberculosis (2001JCM3883), Salmonella spp.

(1996JFP908), Staphylococcus aureus or Yersinia pestis (2009JCM3889).

Bacterial detection using bacteriophages is a reliable and specific method; however,

it requires the genetic engineering of specific phages, which are as yet only available


7

for a few common pathogens. Long assay times are usually required for the

production of detectable luminous signal, especially when few cells of the pathogen

are available.

FMNH2 H3C(CH2)10

O

O2 FMN H2O

H

H3C(CH2)10

OHO

Scheme 1.1: Principle of detection using genetically engineered bacteriophage and enzymatic reaction resulting in bioluminescence (adapted from 2002FRI863).

6

luciferase

BACTERIUM

PHAGE

lux GENE

STEP 1

PHAGE FIXATION

STEP 2

GENE TRANSDUCTION

STEP 3

GENE EXPRESSION

STEP 4

DETECTION via

BIOLUMINESCENCE


8

1.1.2.3 Enzyme-linked immunosorbent assay (ELISA)

Immunoassay utilises the specific relationship between antibodies and antigen

(1990MI497). The specificity and sensitivity of immunoassay has been enhanced by

the use of monoclonal antibodies (specific to one particular type of bacterial antigen)

and the combination of enzyme labels and substrates (1990MI497).

The most popular and widely used type of immunoassay is probably enzyme-linked

immunosorbent assay (ELISA). The principle of ELISA, illustrated in Scheme 1.2,

uses microplate wells coated with capture antibodies.

O OOP

O

OHHO

O OHO

CH3 CH3

After incubation of the bacteria and capture of the antigens, enzyme-linked

antibodies are added to the wells and bind to the fixed antigen (Scheme 1.2). A

fluorogenic enzyme substrate is finally added and, if the correct antigens are present

Scheme 1.2: Bacterial detection using sandwich ELISA (Last accessed 23/03/2010 and adapted from:

http://64.202.120.86/upload/image/articles/2006/biopen/biopen-elisa-schematic.jpg).

Bacterial cell

suspension

- Addition of

enzyme substrate

- Incubation

- Addition of enzyme

linked antibodies

- Incubation

- Washing

- Incubation

- Washing

Capture antibody

Polymer support

Antigens

Enzyme (e.g.: phosphatase)


9

in the resulting ELISA sandwich complex, further incubation results in the detection of

fluorescence.

The most sensitive enzymes used with ELISA are horseradish peroxidase,

phosphatase and β-galactosidase (2005ABI227). This technique has been applied to

the detection of L. monocytogenes (1992LAM26), Ps. fluorescens (1993JAB394), E.

coli O157:H7 (2009BBE1641) or Salmonella spp. (2009BBE1641).

1.1.2.4 Chromogenic media

Chromogenic media utilise enzymes as taxonomic markers.

1.1.2.4.1 Agar-based media

Chromogenic agar-based media are an evolution of the traditional, general purpose,

growth media used in microbiology, which only allow a presumptive identification of

the microorganisms based on their colonial appearance (e.g. pigmentation,

morphology) and require the use of further biochemical test for definitive identification

(2007JAM2046).

The addition of chromogenic enzyme substrates

in those media has allowed a more direct

identification of the suspected pathogen, via the

evidence of a specific enzymatic activity. As

chromogenic media are destined to be used

directly on clinical samples, a combination of

enzyme substrates is often used to differentiate

the pathogen from other commensal bacteria, or

even to allow the identification of several

pathogens simultaneously. Figure 1.2 depicts

the example of a chromogenic medium

(chromID CPS3, bioMérieux) allowing the

simultaneous detection of the most common

urinary tract pathogens: Escherichia coli

(purple), detected via β-glucuronidase activity,

Proteus mirabilis (brown), detected via deaminase activity and enterococci (blue),

detected via β-glucosidase activity. More detail will be given later on the exact nature

of such enzyme substrates. These media also tend to promote the development of

Figure 1.2: Example of a chromogenic medium (chromID CPS3) allowing multiple detection: here E. coli (purple), P. mirabilis (brown) and enterococci (blue). (Picture taken from http://www.biomerieux.fr/servlet/srt/bio/france/dynPage?open=FRN_CLN_PRD&doc=FRN_CLN_PRD_G_PRD_CLN_20&pubparams.sform=2&lang=fr, last accessed 04/04/2010).


10

the microorganism under investigation and the expression of the enzyme necessary

to hydrolyse the substrate, while inhibiting the growth of competing microorganisms,

to facilitate identification and avoid false-positive results (2007BSM96). Thus

chromogenic media can by-pass the time consuming procedure of isolating a pure

culture prior to carrying out further identification tests.

Many media have been developed for the identification of specific pathogens

(2007JAM2046, 2009JMM139), including the detection of antibiotic-resistant

pathogens such as vancomycin-resistant enterococci (VRE, 2009JCM4113,

2009JMM124) and MRSA (2004JCM4519, 2010JCM215).

Chromogenic media confer the advantage to permit procedures and readings free

from any specific and costly equipment.

1.1.2.4.2 Micro gallery and automated system

Micro galleries, such as API gallery (1988AEM2838, 1993ACB81), combine a wide

range of enzyme substrates. A gallery can correctly identify a microorganism within a

given family or genera for which the gallery has been designed. This process has

been automated recently, with systems such as VITEK®2 (bioMérieux), which only

require the manual preparation of the

inoculum. Inoculation of the wells,

incubation and interpretation of the

results are carried out by the system

itself. Figure 1.3 depicts the type of

card used in VITEK®2 system after

incubation, with a card designed for

Gram positive bacteria, showing

positive results corresponding to

presence of Staphylococcus

epidermis. Tests are available for the identification of Enterobacteriaceae

(2003JCM2096), yeasts (2007JCM1087) or Bacillus species (2010LAM120).

The use of galleries allows for the identification of a much wider range of

microorganisms, compared to the agar-based media mentioned previously; however

these techniques of identification usually require the isolation of a pure culture, which

implies longer identification procedures. Automated systems are costly and can only

accommodate a limited number of samples at a time.

Figure 1.3: Example of a VITEK®2 card displaying results for St. epidermis.


11

1.1.3 Chromogenic and fluorogenic dyes

Chromogenic and fluorogenic dyes have been implemented in many detection

techniques, to enhance the visualisation of results by providing a strong and easily

identifiable signal.

They have been divided into 4 classes (B-1980MI05, 2007BSM96) based on their

mode of interaction with microorganisms: DNA fluorescent dyes, pH indicators, redox

indicators and enzyme substrates.

1.1.3.1 DNA fluorescent dyes

DNA fluorescent dyes typically have a low intrinsic fluorescence upon excitation;

however, the electronic alteration resulting from interaction with DNA base pairs

results in a dramatic increase of the quantum yield for the resulting DNA-fluorogen

complex (B-2006MI06). The nature of the interaction is for most dyes, non-covalent,

intercalative binding. DNA fluorescent dyes are usually cationic aromatic

heterocycles, with a planar structure facilitating insertion between the stacked base

pairs of the DNA duplex (B-2006MI06).

Acridine orange 7 (Figure 1.4) is perhaps one of the oldest examples, known to have

two distinct λem whether it is bound to single stranded RNA (λem= 650nm) or double

stranded DNA (λem= 526nm). Phenanthridium dyes 8a-b, mentioned above in section

1.1.1.1 and section 1.1.2.1, are widely used DNA stains; their membrane permeability

can be tuned by modification of the nitrogen substituent (8a-b, Figure 1.4).

8a,R = CH2CH3 λex/em = 510 nm / 595 nm 8b, R = (CH2)3-N

+-(CH3)(CH2CH3)2 λex/em = 530 nm / 625 nm

9, λex/em = 509 nm / 527 nm

7

λex/em = 500 nm / 526 nm (DNA)

Figure 1.4: Examples of some common DNA fluorescent dyes.


12

Thiazole orange 9, an old photographic dye belonging to the class of cyanine dyes,

has shown excellent nucleic acid binding properties associated with an important

increase of its quantum yield upon excitation. This discovery led the preparation of

many analogues such as the SYBR Green I 5 or SYTO dyes (1995US5436134),

mentioned in section 1.1.1.1 and section 1.1.2.1.

These dyes have gained much popularity due to their implementation into modern

detection techniques, which are not limited to microorganism identification, such as

PCR, DNA probes or epifluorescence microscopy to enhance the visualisation of

genetic material or the microorganism itself.

1.1.3.2 pH indicators

Bacterial growth often results in biochemical changes within the growth medium,

some resulting in a noticeable variation of the pH. The addition of a pH indicator to

the growth medium is often used to monitor the production of acid or the release of a

base.

A significant increase of the pH in the growth medium is characteristic of urease

activity or of L-amino-oxidase (deaminase) activity, such as phenylalanine

deaminase, which are both characterised by the release of ammonia. A common

biochemical test used for the detection of these enzymes is the addition of phenol red

(10, Scheme 1.3) into the medium, which results in a change of the medium colour

from yellow to red-fuschia (Scheme 1.3).

A decrease in pH is usually characteristic of sugar fermentation, which results in the

production of lactic acid. MacConkey agar uses neutral red 11 to specifically identify

bacteria that ferment lactose to lactic acid (Scheme 1.4, pH transition: 6.8-8.0).

Scheme 1.3: Acidic and basic forms of phenol red.

10, pH < 6.8 : yellow pH > 8.4 : fuschia


13

1.1.3.3 Redox indicators

Redox indicators detect oxidative enzyme systems, present in all living organisms, by

acting as artificial electron acceptors (2002MI63). They present little specificity and

are used as general bacterial growth indicators.

Methylene blue 12 has been widely used for this purpose, with micro organism

growth monitored by the disappearance of the blue colour resulting from the

formation of leucomethylene blue 13 (Scheme 1.5).

Alternatively, 2,3,5-triphenyltetrazolium chloride (TTC, 14, Scheme 1.6) is also

commonly used as a growth indicator (1987JAB551, 2002MI63) and for the

enumeration of bacterial colonies. TTC is a colourless water soluble salt which forms,

upon reduction, 1,3,5-triphenylformazan 15, a red insoluble solid, which allows clear

visualisation of bacterial development.

Scheme 1.5: Reduction of methylene blue to the colourless leucomethylene blue.

12, Blue

pH < 6.8 : red 11, pH > 8.0: yellow

Scheme 1.4: Acidic and basic forms of neutral red.

Scheme 1.6: Formation of insoluble 1,3,5-triphenylformazan upon reduction of 2,3,5-triphenyltetrazolium chloride.

13, Colourless

15, Red 14, Colourless


14

The ability of 2,3,5-triphenyltetrazolium chlorides to form 1,3,5-triphenylformazan

upon reduction has been widely exploited as a marker of biological activity; the

production of a range of colours can be achieved by substitution or replacement of

the phenyl ring with various heterocyclic moieties.

1.1.3.4 Enzyme substrates

An enzyme substrate is composed of two moieties: a chromophore/fluorophore, or a

precursor susceptible to form one of the former upon a further reaction taking place

after enzymatic hydrolysis, and a biological molecule, usually a peptide or a sugar.

Enzyme substrate dyes comprise an auxochrome, a substituent which enables

linkage to the biological molecule and the formation of an intense colour via electron

conjugation, chelation with a metal, or reaction with a secondary reagent (B-

2008MI07). The two main types of auxochromic groups encountered are the hydroxyl

group, which allows for the detection of osidase, esterase, phosphatase and

sulfatase activities, or the amino group which permits the detection of

aminopeptidase activities.

The efficiency of enzyme substrates has been quantified by the following features:

- The absorption maximum of the free chromophore/fluorophore must be

significantly different from that of the substrate. Ideally the enzymatic substrate will be

non-coloured and/or non U.V. active to allow clear identification of enzymatic activity;

- The chromophore/fluorophore must be readily cleaved and must dissociate

from the enzyme to avoid any interference with the enzymatic activity.

- The chromophore/fluorophore must have a high molar absorptivity to allow

detection of weak enzyme activity;

- The chromophore/fluorophore and its corresponding substrate should be of a

low toxicity to the microorganism(s) under investigation;

- The chromophore/fluorophore and its corresponding substrate should be

stable to the conditions used for bacterial growth: the colour/fluorescence must

develop only upon enzymatic action. Moreover, the chromophore/fluorophore itself

should not undergo any further modification induced by side enzymatic reaction that

would attenuate its colour/fluorescence;

- The substrate should have a good water solubility to ease its incorporation

into the medium and favour its enzymatic hydrolysis;


15

-The chromophore/fluorophore should accurately localise to the bacterial

colony with little or no diffusion of the colour when used in a solid medium.

The overall performances of an enzyme substrate in bacterial identification will be

discussed in terms of sensitivity and specificity; these two criteria are determined by

screening the substrate against a large number of bacterial strains. The sensitivity of

a chromogenic medium is defined as the percentage of positive results for the overall

strains of bacteria expressing the enzyme activity. The specificity is defined as the

percentage of organisms producing a colour being the actual targeted bacterial

strains.

Enzyme substrates have been subdivided into four groups by virtue of their mode of

action (B-1980MI02): simultaneous capture chromogenic substrates, post-incubation

coupling chromogenic substrates, intra-molecular rearrangement/electron

conjugation chromogenic substrates and self-coloured chromogenic/fluorogenic

substrates. Examples of traditional and recently developed enzyme substrates follow

to illustrate this classification.

1.1.3.4.1 Simultaneous capture chromogenic substrates

The product resulting from enzymatic hydrolysis of the substrate, called the primary

reaction product, is further reacted (capture reaction) with a second reagent present

in the medium to form a highly coloured product, called the final reaction product.

Capture reaction has been widely applied to metal chelators such as esculetin

(1987MI188, 16), cyclohexenoesculetin (1996AEM3868, 1997JAM532,

1999US6008008, CHE, 17), 8-hydroxyquinoline (1987MI410, 18), alizarin

(2000LAM336, 2006US7052863, 19), 3-hydroxyflavone (2007MI410, 20) and 3',4'-

dihydroxyflavone (2007MI410, 21, Figure 1.5).

Figure 1.5: Structures of metal chelators used as simultaneous capture chromogenic substrates.

17 19 18

21 20 22


16

Ferric ion, usually incorporated as ferric citrate in the medium, is the metal ion of

choice due to its low toxicity toward microorganisms. The resulting iron complexes

are black for all ligands depicted in Figure 1.5, excepted alizarin, for which the

resulting iron complex is bright violet (2000LAM336).

The capture reaction using ferric ion is depicted in Scheme 1.7 with the example of

β-D-glycosidic derivatives of cyclohexenoesculetin 23 (1996AEM3868, 1997JAM532,

1999US6008008). Upon enzymatic hydrolysis, chelation of the primary reaction

product, cyclohexenoesculetin (CHE, 18), with ferric ions, results in the formation of

the black iron complex 24 (2007BMC1172, Scheme 1.7). The resulting iron complex

24 precipitates out of the medium and locates, with great accuracy, the site of

hydrolysis (Figure 1.6).

The accurate location of the site of hydrolysis, and hence the bacterial colonies,

greatly depends upon the diffusion factor of the primary reaction product and the rate

constant of the capture reaction to form the final reaction product.

Scheme 1.7: Enzymatic hydrolysis of CHE-β-Glu 23, followed by chelation of CHE with ferric ion with Figure 1.6 illustrating the formation of 24 for the detection of Clostridium NC1MB 10697.

24, black complex

18, colourless, blue fluorescence

β-Glucosidase

CHE-β-Glu, 23, colourless solid

Figure 1.6


17

This issue is well illustrated while comparing esculin 17 and CHE 18 (Figure 1.7), the

presence of an extra cyclic carbon chain confers CHE with an increased lipophilicity

resulting in low diffusion within the medium (Figure 1.7).

An alternative to metal chelators in simultaneous capture chromogenic substrates is

the coupling of the primary reaction product to a secondary organic reagent. The

NADI reaction (Scheme 1.8), reported by Ehrlich in 1885 (B-1885MI08), was

appropriately adapted to simultaneous capture reactions.

This reaction was initially used to detect the presence of

cytochrome c oxidase (1954JBC733) via the oxidative

addition of N,N-dimethyl-p-phenylenediamine 25 and α-

naphthol 26, resulting in the formation of the blue

coloured indophenol 27 (Scheme 1.8, Figure 1.6).

Appropriate derivatisation of N,N-dimethyl-p-

phenylenediamine, or an analogous compound, and α-

naphthol have rendered possible the detection of osidase

and amidase microbial activities via this reaction (2002US6340573B1). The most

Figure 1.8: Formation of blue indophenol on agar medium with Pseudomonas aeruginosa.

Scheme 1.8: Formation of the blue indophenol 24 upon enzymatic catalysis

25 26

27, blue

Figure 1.7: Detection of Listeria monocytogenes on agar medium using esculetin-β-D-glucoside (esculin, left) and CHE-β-D-glucoside (right) (L-2008MI01).

[O]


18

advanced example reported allows the detection of up to three different enzyme

activities with the two enzyme substrates L-alanyl-4-amino-2,6-dichlorophenyl-β-D-

galactopyranoside 28 and α-naphthyl-β-D-glucopyranoside 29 (2002US6340573B1,

Scheme 1.9). Formation of the indodichlorophenol dye 32 only occurred upon full

enzymatic hydrolysis and formation of both primary reaction products, 2,6-dichloro-4-

aminophenol 30 and α-naphthol 31 (Scheme 1.9) simultaneously, allowing high

selectivity amongst bacteria.

This reaction is advantageous in terms of specificity compared to metal chelators;

however, it shows less affinity with the site of hydrolysis on solid media where metal

complexes are more accurate, and is best suited for liquid media.

The final and most popular example of this type of substrate are indoxyl derivatives

33-37 (1961T236, 1961JMC574, 1988CJM690);

these substrates are without contest the most

widely used chromogens for the detection of

enzymatic activity and are currently implemented

in most commercially available chromogenic

media (2009JMM139).

Scheme 1.10 illustrates the example of 5-

bromo-6-chloroindoxy-α-glucoside 38, which,

upon hydrolysis of the α-glucoside moiety, releases the free indoxyl 33,

Scheme 1.9: Detection of three different enzyme activities: L-alanine-aminopeptidase and β-galactosidase activities to hydrolyse 28 and β-glucosidase activity to hydrolyse 29.

28

29

32

30

31

NH

HO

R3

R2

R1

33, R1 = H, R2 = Br, R3 = Cl, ‘Magenta’ 34, R1 = Cl, R2 = Br, R3 = H, turquoise (‘X’) 35, R1 = R2 = H, R3 = Cl, ‘Rose/Salmon’ 36, R1 = R3 = H, R2 = Br, blue 37, R1 = R2 = R3 = H, indigo blue


19

spontaneously oxidised to a brightly coloured mixture of indigo 39 and other

indigogenic by-products in minor quantities, such as indirubin 40 (Scheme 1.10). In

this instance, this could be termed a simultaneous self-capture reaction. The resulting

chromogenic mixture formed is highly insoluble and precipitates out of the medium,

allowing a very clear identification of α-glucosidase expressing bacteria (Figure 1.9).

Indoxyls 33-37 are very sensitive substrates with a low toxicity (2007JAM2046); they

are suitable for the detection of osidases, esterases and phosphatases. Moreover the

position and nature of the halogen substituent on the indoxyl moiety offers a large

range of colours, which explains their popularity in microbial detection. Their

synthesis is, however, difficult to achieve and their use is limited to aerobic

microorganisms, as the presence of oxygen is necessary to the formation of the final

reaction product.

1.1.3.4.2 Post-incubation coupling chromogenic substrates

The conditions necessary for the coupling reaction, when using post-incubation

coupling chromogenic substrates, require addition of the secondary reagent after

enzyme hydrolysis, as the growth conditions are strongly affected by the reaction.

Scheme 1.10 and Figure 1.9: Enzymatic hydrolysis of 5-bromo-6-chloroindoxyl (Magenta) α-glucoside and formation of the brightly coloured indigo and indirubin derivatives.

α-glucosidase

[O]

38 33

39

40


20

This method has been mostly applied to azo-coupling reactions, where the secondary

reagent, a diazonium salt, is toxic to microorganisms and inhibitory toward enzymatic

activities (B-1980MI02).

The detection of pyroglutamyl amidase with the substrate pyroglutamyl-β-

naphthylamide 41 is a good illustration of this method: the primary reaction product,

naphthylamine 42, can be coupled with tetrazotized o-anisidine (43, Fast blue B),

resulting in the formation of the blue azo dye 44 (Scheme 1.11, B-1980MI02).

HN

NH

O

pyroglutamylaminopeptidase NH2

NO

pH 1.0 N

N

O

NH2

N

N

N

N

Cl

Cl

H3CO

OCH3 N NN N

H3CO OCH3

NH2 H2N

An alternative capture reaction, using an acidic solution of p-

dimethylaminocinnamaldehyde 45 to form the highly coloured red ene-imine product

46 is also possible (Scheme 1.11, 1987JCM1805, 1996JCM1811).

In both cases the toxicity of the final reaction product forbids any further assay on the

microorganism under investigation, and this resulted in research on other, less toxic,

substrates (1967MI500, 2002US6340573B1).

Scheme 1.11: Enzymatic hydrolysis of pyroglutamyl-β-naphthylamide 41 and capture of the resulting product, β-naphthylamine 42.

41 42

43

44, blue

45

46, red/ red-violet

42


21

More recently, the example of 9-(4'-aminophenyl)acridine substrates was introduced

as a means of detection for peptidase activities (2007BMCL1418). Although not a

true coupling reaction, the primary reaction product 47 is weakly coloured and

requires the addition of acetic acid to yield the colourful 9-(4'-aminophenyl)acridinium

ion (48, Scheme 1.12, Figure 1.10, 2007BMCL1418).

Acidification of the medium is only possible after incubation; as such a low pH is

inhibitory to the growth of most bacteria.

Post-incubation coupling chromogenic substrates are now seldom used; the

important range of other enzyme substrates available now can usually provide a

better alternative to post-coupling reactions.

1.1.3.4.3 Intramolecular rearrangement/electron conjugation chromogenic

substrates

In this instance, the enzyme induces a structural modification, such as an

intramolecular rearrangement, forcing a change of the electron conjugation within the

molecule and a shift in the wavelength of light absorption.

An example of this type of substrate is nitrocefin 47, which is utilised in the detection

of β-lactamase enzyme producing bacteria (2005JOC367). Upon hydrolysis of the

Scheme 1.12: Ionisation of 9-(4'-amino-2',6'-dimethoxyphenyl)acridine with acetic acid and consequent formation of a purple colour (2007BMLC1418). Figure 1.10: Importance of ionisation of 33 for the visualisation of the bacterial colonies.

47 48

Protonated form

Neutral form


22

β-lactam ring in nitrocefin 47 by β-lactamase activity, the altered electron density in

cephalothin 48 induces a dramatic change of the wavelength absorbance, permitting

visible colony detection (Scheme 1.13).

Recently, researchers from Biosynth® have exploited this technique by developing a

range of indoxyl enzyme substrates, such as the ALDOL™-455-β-D-galactosidase 49

(P-2009MI01). Upon enzymatic hydrolysis of 49, the colourless primary reaction

product 50 undergoes an intramolecular Aldol-type condensation to produce a yellow

insoluble dye, 7-chloro-10-phenyl-10H-indolo-[1,2a]-indol-10-one 51 (Scheme 1.14).

Scheme 1.13: Opening of the nitrocefin β-lactam ring by β-lactamase enzymatic activity.

47, yellow, λmax: 390nm 48, red, λmax: 486nm

β-lactamase

Scheme 1.14 and Figure 1.11: Intramolecular rearrangement of ALDOL™-455-β-D-galactosidase 49 upon enzymatic hydrolysis and resulting yellow colonies of Enterobacter cloacae (P-2009MI01, with Dr. L.M. Wick’s authorisation).

β-D-galactosidase

49, colourless

51, yellow

50


23

The rearrangement appears to be instantaneous, and the precipitation of the

resulting product 51 locates the bacterial colonies with precision (Figure 1.11). The

low toxicity and absence of background of these substrates is also notable (Figure

1.11).

Very few examples of intramolecular rearrangement substrates are available yet,

presumably due to the difficulty of finding an adequate rearrangement reaction,

suitable to the bacterial growth condition.

1.1.3.4.4 Self coloured chromogenic/fluorogenic substrates

The primary reaction product itself is coloured/fluorescent suppressing the need of

any other reagents.

The colouration/fluorescence of this type of enzyme substrates results from the

conjugation of an available lone pair of electrons, belonging to the auxochrome, into

the π-system of the chromophore/fluorophore. The linkage of a biological molecule to

the auxochromic group translates into a dramatic quenching of the

colour/fluorescence for the resulting enzyme substrate, as a result of important

changes within the electron conjugation. Scheme 1.15 depicts the electron

movement within the core structure 52 common to some popular chromogenic and

fluorogenic dyes, and the restricted conjugation occurring within the bond formed

between the dye and the biological molecule, in the corresponding enzyme substrate

53. Cleavage of the biological marker by enzymatic hydrolysis restores its initial

colour/fluorescence.

Scheme 1.15: Electron movements and quenching of this effect in some common structure of self-coloured enzyme substrates: X = O or N, X2 = O or C(CH3)2, X

3 = N, C-Ph or C-PhCOOH, R = Alkyl chain or amino acid

52

Coupling

Hydrolysis

Colour/ Fluorescence Quenched colour/ fluorescence

53


24

1.1.3.4.4.1 Fluorogen

The three most common fluorogenic cores (Figure 1.12) used as enzyme substrates

are fluorescein 54, 7-hydroxyphenoxazin-3-one (resorufin, 55) and, probably the

most important class, the coumarin core, including 7-hydroxy-4-methylcoumarin (4-

methylumbelliferone, 4-MU, 56a) and 7-amino-4-methylcoumarin (7-AMC, 56b)

(2009JMM139). Recently, benzoxazole 57a and benzothiazole 57b derivatives have

been used successfully as enzyme substrates (2008WO152306).

.

The most widely used fluorogen is probably 4-MU, partly due to its low toxicity to

microorganisms, ease of preparation and bright blue fluorescence under U.V.

excitation. It is commonly utilised for the detection of microbial sugar hydrolase

activity. However, the use of 4-MU (pKa = 7.80)

has several disadvantages. The pH dependence

is an important issue, as full or partial dissociation

of the phenolic proton is essential to enhance

electron conjugation and reach maximum

fluorescence intensity. This issue has been

addressed by the preparation of several 7-

hydroxycoumarin derivatives possessing a lower

pKa and allowing full dissociation of the phenolic

proton at relatively low pH: fluorinated coumarins

and particularly 6,8-difluoro-4-methylumbelliferone

(1997BMC1985, 1998US5830912) as well as chlorinated coumarin analogues

(2006JAM977), showed higher efficiency and sensitivity than the traditional 4-MU

enzyme substrate. If the full dissociation of the phenolic proton improves the

56a, R = OH, λex/em : 360/449nm 56b,R = NH2, λex/em : 351/430nm

54, λex/em : 494/518nm

55, λex/em : 571/595nm

Figure 1.12: Structure of common fluorogens.

Figure 1.13: Diffusion of 4-MU on a multipoint agar plate.

57a = O 57b = S


25

emission intensity of the fluorogen, it consequently augments its solubility in aqueous

media, which dramatically increases the diffusion of the fluorogen on agar media,

considerably limiting its usefulness in locating bacterial colonies on solid media. As

an example, the screening of organisms via multipoint inoculation is rendered

extremely difficult due to the diffusion of 4-MU in the medium (Figure 1.13).

The relatively low pKa of fluorescein (pKa = 6.4) and of resorufin (pKa = 6.0) results

in partial deprotonation at physiological pH, and

causes the same diffusion phenomenon when

these substrates are used on agar media.

Resorufin-β-D-glucuronide is commonly used for

the detection of E. coli; however, the product of

enzymatic hydrolysis shows poor localisation of

the bacterial colonies (Figure 1.14). Its derivative,

2-dodecylresorufin is reported to show high affinity

for lipid regions of cells (B-2005MI04); however,

its use as an enzyme substrate has not been

reported yet. The preparation of fluorinated resorufin analogues has also been

reported to improve the fluorogenic properties of these dyes (2008US7432372).

Despite the disadvantages mentioned above, fluorogenic substrates are notoriously

very sensitive with diagnostic results achievable within hours. Early readings are not

achievable with the naked eye and require instrumentation to measure weak

fluorescence. Moreover, their use in solid media has been restricted in favour of

chromogenic substrates (2007BSM96).

1.3.4.4.2 Chromogens

Early development of self-coloured enzyme substrates involved the use of p-

nitroaniline 58a (1961ABB271, 1967MI415) and p-nitrophenol

58b (1961BBA460, 1967SCI1451). Enzyme activity was

witnessed by the appearance of a yellow colour at the site of

hydrolysis. Although, the colour is relatively weak compared to

other chromogens, these enzyme substrates are still used in

various test kits, as they are cheap and easy to produce.

More recently, several 7-aminophenoxazin-3-one derivatives 59a-d have been

reported for the detection of aminopeptidase activity on agar medium (2008OBC682).

Figure 1.14: Detection of E. coli using resorufin-β-D-glucuronide.

58a, R = NH2, yellow 58b, R = OH, yellow

NO2

R


26

The presence of a fused benzene ring (2006US0121551), or alkyl substituent

(2006WO030119, 2008OBC682), and the absence of a significant pH dependency,

successfully limited the colour diffusion from the site of hydrolysis in solid media.

7-Amino-1-pentylphenoxazin-3-one 59a is a particularly efficient substrate, with high

affinity for the bacterial colonies (Figure 1.15). The main drawback reported for 7-

aminophenoxazin-3-one based enzyme substrates is the orange background

colouration generated by the substrate itself

(Figure 1.15).

Other, less popular, chromogens were reported for detection of enzymatic activity:

acridinone 60, (1989US4810636, 1991AGE1646), naphtholbenzein 61

(2000AEM5521), 5-(4-hydroxy-3,5-dimethoxyphenylmethylene)-2-thioxothia-zolidin-

4-one-3-ethanoic acid (SRA, 62, 2009FML10) and the 3-methoxy and 3,5-dimethoxy

derivatives of 4-[2-(4-hydroxy-3,5-dimethoxyphenyl)-vinyl]-quinolinium-1-(propan-3-yl

carboxylic acid) (63a [SLPA] and 63-b [VLPA], 1990CARc5, 1999AEM807,

2002AEM3622) proven to be efficient for bacteria identification; colourfull colonies

were formed upon hydrolysis of their corresponding substrates (Figure 1.16).

N

HO O

OHO

OH

R1R2

N

HOOC

OH

OCH3H3CO

S

NO

S

COOH

59a, R1 = n-pentyl, R2 = R3 = H, red/pink 59b, R1 = R2 = CH3, R3 = H, red/pink 59c, R1 = R2 = R3 = CH3, red/pink 59d, R1 = R2 = -(CH)4-, R3 = H, red/pink

Figure 1.15: Detection of Ps .aeruginosa using 7-N-β-alanylamino-1-pentylphenoxazin-3-one.

60, Pink 61, Red-Pink

O

N

H2N O

R1R2

R3

62, Red-Pink

63a, R1 = R2 = OCH3, burgundy 63b, R1 = OCH3, R2 = H, red

Figure 1.16: Structure of chromogens recently developed for the detection of enzymatic activity.


27

1.1.4 Enzyme targets

Knowledge of the most common enzymes expressed by pathogenic and non-

pathogenic microorganisms is essential to the successful development of efficient

enzyme substrates. Identification and characterisation of all the different enzymes

produced by a bacterium is a long and tedious process (2007BSM96) and such

information is not always available. However, a good knowledge of the most

important bacterial enzymes has been acquired. A description of enzyme classes and

their use in microbiology for the differentiation of microorganisms has been given by

Bascomb (1987MM105) and more recently by Orenga et al. (2009JMM139). The

following review will attempt to highlight briefly the main enzyme activities exploited in

chromogenic media for the detection of some important pathogens.

1.1.4.1 Esterases and lipases

Esterase enzymes are ubiquitous in all living organisms. Esterases can hydrolyse

substrates with a short carbon chain, whereas lipase enzymes hydrolyse long carbon

chain esters and trialkylglycerol fatty acid esters to glycerol and the constituent fatty

acids. Hydrolysis of short esterase substrates, such as various fluorescein diacetate

derivatives, has been widely used for the monitoring of microorganism viability and

activity by flow cytometry (1995FML1, 2003JMM379). Variation of the ester

hydrocarbon chain has been exploited to achieve higher specificity, for example, use

of the fluorogenic substrate 4-methylumbelliferyl butyrate has been suggested as a

complementary test for the differentiation of various microorganisms, such as

Branhamella catarrhalis and Neisseria spp. (1988JCM1227) or Mycobacterium

fortuitum and Mycobacterium chelonei (1977MI147).

The most important application of chromogenic subtrates to esterase activity is

probably for the detection in stool samples of Salmonella spp., the pathogen

responsible for the majority of food poisoning episodes in the United Kingdom

(1999JCM766). Salmonella spp. are known to be some of the few

Enterobacteriaceae able to hydrolyse fatty acid esters with carbon chain lengths of C-

7 to C-10 (2007JAM2046). Cooke et al. studied the sensitivity of various esters of 4-

[2-(4-hydroxy-3,5-dimethoxyphenyl)-vinyl]-quinolinium-1-(propan-3-yl carboxylic acid)

bromide (SLPA, 63a), differing in chain length (C-4 to C-10), and found the

octanoate ester to be most sensitive for the detection of Salmonella spp. (64,

Scheme 1.16, 1999AEM807).


28

Consequently, many recent commercial chromogenic substrates rely on the

octanoate esterase activity to detect Salmonella spp.: as an example, indoxyl

magenta caprylate is present in Oxoid-Salmonella-Chromogen-Agar (OSCM, Oxoid),

Rapid'Salmonella (BioRad), Compass Salmonella (Biokar diagnostic), Salmonella-

Agar-Plate (ASAP, AES), Salmonella-Medium-Identification-Detection (SM ID2,

bioMérieux) or HiCrome Salmonella agar (HIMEDIA) (2007BCM96). A comparative

study carried by Perez et al. (2003JCM1130) confirmed the high specificity and

selectivity of such detection media for the elucidation of Salmonella spp. within

clinical specimens.

The lipase enzyme activity has been seldom exploited in the field of enzyme

substrates, presumably due to the difficulty of incorporating highly lipophilic enzyme

substrates into an aqueous medium. The hydrolysis of dialkylglycerol fatty acid esters

of resorufin by a free lipase from Pseudomonas cepacia (2006JMCE76) has,

however, been reported, suggesting potential applications.

1.1.4.2 Glycosidases

1.1.4.2.1 β-D-Glucuronidase

Bacterial β-D-glucuronidase plays a role in the decomposition process of the host

connective tissue during the infectious process (1973AM863).

β-D-Glucuronidase activity is relatively limited amongst bacteria; it has been detected

mainly in Escherichia coli (94 to 96% of clinical isolates), but also in some Shigella

species (44 to 58% of clinical isolates) and Salmonella species (20 to 29% of clinical

isolates) (1991MR335). The prevalence of this enzyme in E. coli (1990AEM1203) has

Salmonella spp.

esterase

Scheme 1.16: Hydrolysis of SLPA octanoate 63 by Salmonella spp. esterase (1999AEM807).

64 63a


29

generated a strong interest in the preparation of β-D-glucuronic acid derivatives: with

chromogens, such as p-nitrophenol (1984JCM1177, 1990AEM2021), 5-bromo-4-

chloroindoxyl (1988CJM690, 1988AEM1874), or fluorogens, such as 4-

methylumbelliferone (1984JFS1186, 1986JCM368, 1988JCM2682) and its 6-chloro

derivative (2006JAM977). These substrates can differentiate with high specificity E.

coli (the most common urinary tract pathogen) amongst other Enterobacteriaceae

present in urine samples (1995JCM199, 2009JMM139). Current commercial

chromogenic agars such as chromID coli (bioMérieux) or Oxoid BrillianceTM

E.coli/coliform Selective Agar (Oxoid) exploit the β-D-glucuronidase acitivity of E. coli.

. 1.1.4.2.2 β-D-Galactosidase

β-D-Galactosidase, also called lactase, catalyses the breakdown of lactose 65 into its

monosaccharide constituents β-D-galactose 66a and α/β-D-glucose 66b (Scheme

1.17), a step involved in the fermentation of sugar. This enzyme is mainly distributed

within the coliform group (Enterobacteriaceae), which are common water pollutants;

assay of β-D-galactosidase activity is therefore part of the national guidelines for the

microbiological examination of water (2001JAM1118).

The detection of β-D-galactosidase activity with chromogenic substrates was first

introduced by Aizawa using o-nitrophenyl-β-D-galactoside (1939MI321). Numerous β-

D-galactopyranosyl substrates have been prepared since then, using p-nitrophenol,

6-bromo-2-naphtol (1967MI395), 5-bromo-4-chloro-3-indoxyl (1990AEM301), alizarin

(2000LAM336), CHE (1996AEM3868), p-naphtholbenzein (2000AEM5521) as

chromogens, and 4-methylumbelliferone (4-MU) or 7-hydroxycoumarin-3-carboxylate

(2001JAM1118) as fluorogens.

Specific chromogenic media for some virulent member of the coliform group such as

E. coli 0157:H7 or vancomycin resistant enterrococci (VRE) are based on a β-D-

galactopyranosyl derivative of indoxyl or alizarin (2009JMM139).

β-Galactosidase

Scheme 1.17: Lactose hydrolysed into β-D-galactose and α/β-D-glucose.

65 66a 66b


30

1.1.4.2.3 α-D-Galactosidase

Little information is available on the prevalence of α-D-glycosidase, however, α-D-

galactopyranoside activity seems to occur in some species of the genus

Streptococcus and Enterococcus (1989JCM1719). Streptoccocus bovis, which is

linked with gastrointestinal neoplasia, was differentiated from other streptococci via

α-D-galactopyranoside activity using 4-MU (1983AEM622) or p-nitrophenol

(1989JCM1719).

Perry et al. reported this enzyme to be predominant in Salmonella spp., and

consequently developed an agar medium (ABC medium, 1999JCM766) including an

α-D-galactoside chromogen, to differentiate Salmonella spp. from the various

Enterobacteriaceae present in stool samples.

1.1.4.2.4 β-D-Glucosidase

The primary role of β-D-glucosidase is the hydrolysis of gluco-oligosaccharides into

single glucose units. This enzyme is widely

distributed amongst bacteria and has found

application for the detection of some important

human pathogens (2009JMM139). β-D-Glucosidase

is prevalent in enteroccoci (1997JAM532,

2006JAM410), Listeria spp. (2004MI1), Vibrio spp.

(2005MI1454), Candida spp. (2009JMM139) and members of the Enterobacteriaceae

family (1997JAM532, 2006JAM410).

The use of the natural substrate esculin 67 for the detection of β-D-glucosidase

activity has been known for more than a century (1909MI547), but the issue of

diffusion discussed in section 1.3.4.1 (Figure 1.7) led to the use of more efficient

substrates. Perry et al. tested most known chromogen for the detection of β-D-

glucosidase activity: indoxyls, 8-hydroxyquinoline, esculetin, CHE, 3-hydroxyflavone,

3',4'-dihydroxyflavone and alizarin, and highlighted the variation of sensitivity of a

same substrate amongst bacterial species and the necessity to select carefully a

chromogen for targeting a particular pathogen (1997JAM532).

67

O O

HO

OHO

HOOH

O

OH


31

1.1.4.2.5 αααα-D-Glucosidase

Sadler et al. demonstrated the occurence of α-D-glucosidase activity in the genus

Bacilli using p-nitrophenyl-α-D-glucoside (1984JCM594) and differentiated the

pathogenic Bacilli anthracis, responsible for the disease commonly called anthrax,

from other non pathogenic Bacilli (1984JCM594).

More recent applications have exploited α-D-glucosidase activity to differentiate

Enterococcus faecalis from Enterococcus faecium (VRE-BMX, bioMérieux,

2007JCM1556), to identify specifically Staphylococcus aureus (S. aureus ID,

bioMérieux, 2003JCM5695), or Chronobacter sakazakii (2007AEM48), an occasional

contaminant of powdered infant formula milk. All media used indoxyl-α-D-glucoside

derivatives as the chromogen.

1.1.4.2.6 β-Hexoaminidase

β-Hexoaminidase enzyme is prevalent mainly in Candida albicans, a commensal

yeast of the digestive mucosa, which can proliferate under certain condition, causing

severe mycoses. This enzyme is expressed weakly, if at all, by other Candida

species (2002AEM3622, 2006JCM3340), and provides the ideal tool for the

differentiation of C. albicans.

Commercially available chromogenic media, such as CHROMagar Candida

(CHROMagar, 1996JCM454), Candida ID 2 (CAID2, bioMérieux, 2006JCM3340),

Candida diagnostic agar (CDA, PPR diagnostic limited, 2002AEM3622) and

CandiSelect4 (CS4, Bio-Rad, 2008JMM89) exploit β-hexoaminidase activity for the

identification of C. albicans. Cooke et al. reported the use of a new substrate,

ammonium 4-{2-[4-(2-acetamido-2-deoxy-β-D-glucopyranosyloxy)-3-methoxyphenyl]-

vinyl}-1-(propan-3-yl-oate)-quinolium bromide (VLPA-GlcNAc, 68, Scheme 1.18,

2002AEM3622), which produced red colonies of C. albicans.


32

Several comparative studies (1996JCM454, 1998JMM623, 2001MI9, 2002AEM3622)

have highlighted the high sensitivity, high specificity and time-saving of such

chromogenic media, suggesting them as an advantageous replacement of

conventional C. albicans tests in microbiological routine tests.

1.1.4.3 Phosphatase

Phosphatase enzymes are involved in several key processes and, hence, are widely

distributed amongst organisms. Their use in the domain of bacterial identification is

limited to the hydrolases of phosphate ester, the latter being divided into two

c

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